Skip to main content

Thank you for visiting nature.com. You are using a browser version with limited support for CSS. To obtain the best experience, we recommend you use a more up to date browser (or turn off compatibility mode in Internet Explorer). In the meantime, to ensure continued support, we are displaying the site without styles and JavaScript.

  • Review Article
  • Published:

The biofilm matrix: multitasking in a shared space

Abstract

The biofilm matrix can be considered to be a shared space for the encased microbial cells, comprising a wide variety of extracellular polymeric substances (EPS), such as polysaccharides, proteins, amyloids, lipids and extracellular DNA (eDNA), as well as membrane vesicles and humic-like microbially derived refractory substances. EPS are dynamic in space and time and their components interact in complex ways, fulfilling various functions: to stabilize the matrix, acquire nutrients, retain and protect eDNA or exoenzymes, or offer sorption sites for ions and hydrophobic substances. The retention of exoenzymes effectively renders the biofilm matrix an external digestion system influencing the global turnover of biopolymers, considering the ubiquitous relevance of biofilms. Physico-chemical and biological interactions and environmental conditions enable biofilm systems to morph into films, microcolonies and macrocolonies, films, ridges, ripples, columns, pellicles, bubbles, mushrooms and suspended aggregates — in response to the very diverse conditions confronting a particular biofilm community. Assembly and dynamics of the matrix are mostly coordinated by secondary messengers, signalling molecules or small RNAs, in both medically relevant and environmental biofilms. Fully deciphering how bacteria provide structure to the matrix, and thus facilitate and benefit from extracellular reactions, remains the challenge for future biofilm research.

This is a preview of subscription content, access via your institution

Access options

Buy this article

Prices may be subject to local taxes which are calculated during checkout

Fig. 1: The matrix of microbial biofilms.
Fig. 2: Interactions of extracellular polymeric substance components and their functions.
Fig. 3: Regulation of matrix synthesis.
Fig. 4: Mechanical properties of the biofilm matrix.

Similar content being viewed by others

References

  1. Wingender, J., Neu, T. R. & Flemming, H.-C. in Microbial Extracellular Polymeric Substances (eds. Wingender, J., Neu, T. R. & Flemming, H.-C.) 1–19 (Springer, 1999).

  2. Steinberg, N. & Kolodkin-Gal, I. The matrix reloaded: how sensing the extracellular matrix synchronizes bacterial communities. J. Bacteriol. 197, 2092–2103 (2015).

    Article  CAS  Google Scholar 

  3. Frantz, C., Stewart, K. M. & Weaver, V. M. The extracellular matrix at a glance. J. Cell Sci. 123, 4195–4200 (2010).

    Article  CAS  Google Scholar 

  4. Karygianni, I., Ren, Z. & Thurnheer, T. Biofilm matrixome: extracellular components in structured microbial communities. Trends Microbiol. 28, 668–681 (2020).

    Article  CAS  Google Scholar 

  5. Serra D. O. & Hengge, R. in Extracellular Sugar-Based Biopolymers Matrices. Biologically-Inspired Systems Vol. 12 (eds Cohen, E. & Merzendorfer, H.) 355–392 (Springer, 2019).

  6. Dueholm, M. S. & Nielsen, P. H. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 113–133 (IWA, 2017).

  7. Erskine, E. et al. Formation of functional, non-amyloidogenic fibres by recombinant Bacillus subtilis TasA. Mol. Microbiol. 110, 897–913 (2018).

    Article  CAS  Google Scholar 

  8. Neu, T. R. & Lawrence, J. R. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 25–60 (IWA, 2017.

  9. Toyofuku, M., Nomura, N. & Eberl, L. Types and origins of bacterial membrane vesicles. Nat. Rev. Microbiol. 17, 13–24 (2019). This work presents a comprehensive and systematic overview of OMVs, outer–inner membrane vesicles and cytoplasmatic membrane vesicles and their formation and functions.

    Article  CAS  Google Scholar 

  10. Choi, S. Y. et al. Chromobacterium violaceum delivers violacein, a hydrophobic antibiotic, to other microbes in membrane vesicles. Environ. Microbiol. 22, 705–713 (2020).

    Article  CAS  Google Scholar 

  11. Wurl, O. & Cunliffe, M. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 249–268 (IWA, 2017).

  12. Flemming, H.-C. et al. Biofilms: an emergent form of microbial life. Nat. Rev. Microbiol. 14, 563–575 (2016).

    Article  CAS  Google Scholar 

  13. Jennings, L. K. et al. Pseudomonas aeruginosa aggregates in cystic fibrosis sputum produce exopolysaccharides that likely impede current therapies. Cell Rep. 34, 108782 (2021).

    Article  CAS  Google Scholar 

  14. Costerton, J. W. et al. Bacterial biofilms in nature and disease. Annu. Rev. Microbiol. 41, 435–464 (1987).

    Article  CAS  Google Scholar 

  15. Hengge, R. Linking bacterial growth, survival and multicellularity — small signaling molecules as triggers and drivers. Curr. Opin. Microbiol. 55, 57–66 (2020). This paper shows clearly how the transition to multicellularity is achieved by a regulatory signalling network, promoting either growth or stress resistance, and how the matrix represents a self-constructed homeostatic ‘niche’.

    Article  CAS  Google Scholar 

  16. Penesyan, A., Paulsen, I. T., Kjelleberg, S. & Gillings, M. R. Three faces of biofilms: a microbial lifestyle, a nascent multicellular organism, and an incubator for diversity. NPJ Biofilms Microbiomes 7, 80 (2021).

    Article  CAS  Google Scholar 

  17. Dragoš, A. et al. Division of labour during biofilm matrix production. Curr. Biol. 28, 1903–1913 (2018).

    Article  Google Scholar 

  18. Decho, A. W. & Guiterrez, T. Microbial extracellular substances (EPSs) in ocean systems. Front. Microbiol. 8, 922 (2017).

    Article  Google Scholar 

  19. Oppenheimer-Shaanan, Y. et al. Spatio-temporal assembly of functional mineral scaffolds within microbial biofilms. NPJ Biofilms Microbiomes 2, 15031 (2016).

    Article  Google Scholar 

  20. Staudt, C., Horn, H., Hempel, D. C. & Neu, T. R. in Biofilms in Medicine, Industry and Environmental Technology (eds Lens, P. et al.) 308–327 (IWA, 2003).

  21. Bennke, C. M., Neu, T. R., Fuchs, B. M. & Amann, R. Mapping glycoconjugate-mediated interactions of marine Bacteroidetes with diatoms. Syst. Appl. Microbiol. 36, 417–425 (2013).

    Article  CAS  Google Scholar 

  22. Maqbool, T., Cho, J., Shin, K. H. & Hur, J. Using stable isotope labeling approach and two dimensional correlation spectroscopy to explore the turnover cycles of different carbon structures in extracellular polymeric substances. Water Res. 170, 115355 (2020).

    Article  CAS  Google Scholar 

  23. Choong, F. X. et al. Real-time optotracing of curli and cellulose in live Salmonella biofilms using luminescent oligothiophenes. NPJ Biofilms Microbiomes 2, 16024 (2016).

    Article  Google Scholar 

  24. Seviour, T. W. et al. Extracellular polymeric substances of biofilms: suffering from an identity crisis. Water Res. 151, 1–7 (2019). This work presents a wide spectrum of EPS components, and an inspiring multidisciplinary road map for addressing the nature, function and control of these components is proposed.

    Article  CAS  Google Scholar 

  25. Flemming, H.-C. & Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 8, 623–633 (2010).

    Article  CAS  Google Scholar 

  26. Bruckner, C. G., Rehm, C., Grossart, H. P. & Kroth, P. G. Growth and release of extracellular organic compounds by benthic diatoms depend on interactions with bacteria. Environ. Microbiol. 13, 1052–1063 (2011).

    Article  CAS  Google Scholar 

  27. Pierce, C. et al. The Candida albicans biofilm matrix: composition, structure and function. J. Fungi 3, 14 (2017).

    Article  Google Scholar 

  28. Tomer, A. et al. in Mycoremediation and Environmental Sustainability. Fungal Biology (eds Prasad, R., Nayak, S. C., Kharwar, R. N. & Dubey, N. K.) 187–200 (Springer, 2021).

  29. Turnheer, T., Gmür, R., Shapiro, S. & Guggenheim, B. Mass transport of macromolecules within an in vitro model of supragingival plaque. Appl. Environ. Microbiol. 60, 1702–1709 (2003).

    Article  Google Scholar 

  30. Boles, B. R. & Horswill, A. R. Swimming cells promote a dynamic environment within biofilms. Proc. Natl Acad. Sci. USA 109, 32 (2012).

    Article  Google Scholar 

  31. Piard, J. C. et al. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 179–191 (IWA, 2017).

  32. Costerton, J. W., Geesey, G. G. & Cheng, K.-J. How bacteria stick. Sci. Am. 238, 86–95 (1978).

    Article  CAS  Google Scholar 

  33. Turnbull, J. E. & Field, R. A. Emerging glycomics technologies. Nat. Chem. Biol. 3, 74–77 (2007).

    Article  CAS  Google Scholar 

  34. Sutherland, I. W. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 15–24 (IWA, 2017).

  35. Limoli, D. H. et al. Bacterial extracellular polysaccharides in biofilm formation and function. Microbiol. Spectr. https://doi.org/10.1128/microbiolspec.MB-0011-2014 (2015). This work presents a good overview of the aggregative, structural and protective properties of polysaccharides in the matrix which provide the successful adaptation of bacteria to nearly every niche.

    Article  Google Scholar 

  36. Pestrak, M. J., Eggleston, H. C. & Wozniak, D. J. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 79–112 (IWA, 2017). This work is a good overview of P. aeruginosa polysaccharides, and their composition, structure, biosynthesis, functions and regulation.

  37. Tseng, B. S. et al. A biofilm matrix-associated protease inhibitor protects Pseudomonas aeruginosa from proteolytic attack. mBio 9, e00543-18 (2018).

    Article  Google Scholar 

  38. Whitfield, C., Wear, S. S. & Sande, C. Assembly of bacterial capsular polysaccharides and exopolysaccharides. Annu. Rev. Microbiol. 74, 521–543 (2020).

    Article  CAS  Google Scholar 

  39. Abdian, P. L. & Zorreguieta, A. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R., Wingender, J.) 227–247 (IWA, 2017).

  40. Caudan, C., Filali, A., Spérandio, M. & Girbal-Neuhauser, E. Multiple EPS interactions involved in the cohesion and structure of aerobic granules. Chemosphere 117, 262–270 (2014).

    Article  Google Scholar 

  41. Felz, S., Neu, T. R., van Loosdrecht, M. C. M. & Lin, Y. Aerobic granular sludge contains hyaluronic acid-like and sulfated glycosaminoglycan-like polymers. Water Res. 169, 115291 (2020).

    Article  CAS  Google Scholar 

  42. de Graaff, D. R. et al. Sialic acids in the extracellular polymeric substances of seawater-adapted aerobic granular sludge. Wat. Res. 155, 343–351 (2019).

    Article  Google Scholar 

  43. Bonnans, C., Chou, J. & Werb, Z. Remodelling the extracellular matrix in development and disease. Nat. Rev. Mol. Cell Biol. 15, 786–801 (2014).

    Article  CAS  Google Scholar 

  44. Brown, A. J. XLIII.— On an acetic ferment which forms cellulose. J. Chem. Soc. Trans. 49, 432–439 (1886).

    Article  CAS  Google Scholar 

  45. Römling, U. & Galperin, M. Y. Bacterial cellulose biosynthesis: diversity of operons, subunits, products, and functions. Trends Microbiol. 23, 545–557 (2015).

    Article  Google Scholar 

  46. Yamananka, S. et al. The structure and mechanical properties of sheets prepared from bacterial cellulose. J. Mat. Sci. 24, 3141–3145 (1989).

    Article  Google Scholar 

  47. Ziemba, C., Shabtai, Y., Piatkovsky, M. & Herzberg, M. Cellulose effects on morphology and elasticity of Vibrio fischeri biofilms. NPJ Biofilms Microbiomes 2, 1 (2016).

    Article  Google Scholar 

  48. Thongsomboon, W. et al. Phosphoethanolamine cellulose: a naturally produced chemically modified cellulose. Science 359, 6373 (2018).

    Article  Google Scholar 

  49. Grossart, H.-P., Tang, K. M., Kiørboe, T. & Ploug, H. Comparison of cell-specific activity between free-living and attached bacteria using isolates and natural assemblages. FEMS Microbiol. Lett. 266, 194–200 (2007).

    Article  CAS  Google Scholar 

  50. Wingender, J. & Jaeger, K. E. in Encyclopedia of Environmental Microbiology (ed. Bitton, G.). 1207–1223 (Wiley, 2002).

  51. Zhang, P. et al. Identification and function of extracellular protein in wastewater treatment using proteomic approaches: a minireview. J. Environ. Manag. 233, 24–29 (2019).

    Article  CAS  Google Scholar 

  52. Tielen, P. et al. Interaction between extracellular lipase LipA and the polysaccharide alginate of Pseudomonas aeruginosa. BMC Microbiol. 13, 159 (2013).

    Article  CAS  Google Scholar 

  53. Li, Q. & Sand, W. Mechanical and chemical studies on EPS from Sulfobacillus thermosulfidooxidans: from planktonic to biofilm cells. Coll. Surf. B Biointerfaces 153, 34–40 (2017).

    Article  CAS  Google Scholar 

  54. Lindsay, S., Oates, A. & Bourdillon, K. The detrimental impact of extracellular proteases on wound healing. Int. Wound J. 14, 1237–1247 (2017).

    Article  Google Scholar 

  55. McDougald, D., Rice, S. A., Barraud, N., Steinberg, P. D. & Kjelleberg, S. A. Should we stay or should we go: mechanics and ecological consequences of biofilm dispersal. Nat. Rev. Microbiol. 10, 39–50 (2012). This classic work on the reasons for and mechanisms of biofilm dispersal shows that the disassembly of biofilms is as equally regulated as their formation.

    Article  CAS  Google Scholar 

  56. Rumbaugh, K. P. & Sauer, K. Biofilm dispersion. Nat. Rev. Microbiol. 18, 571–581 (2020). This work highlights the role of differentiated dispersal cells and develops a broad conceptual framework for the diversity of mechanisms leading to biofilm disassembly, portraying dispersal as an active event in which biofilm cells convert to the planktonic mode of growth.

    Article  CAS  Google Scholar 

  57. Flemming, H.-C. & Wuertz, S. Bacteria and archaea on Earth and their abundance in biofilms. Nat. Rev. Microbiol. 17, 247–260 (2019).

    Article  CAS  Google Scholar 

  58. Falkowski, P. G., Fenchel, T. & Delong, E. F. The microbial engines that drive Earth’s biogeochemical cycles. Science 320, 1034 (2008).

    Article  CAS  Google Scholar 

  59. Battin, T. J., Besemer, K., Bengtsson, M. M., Romani, A. M. & Packmann, A. I. The ecology and biogeochemistry of stream biofilms. Nat. Rev. Microbiol. 14, 251–263 (2016). This work demonstrates the extent to which biofilms dominate microbial life in streams and rivers, drive crucial ecosystem processes and contribute to global biogeochemical fluxes.

    Article  CAS  Google Scholar 

  60. Álvarez-Mena, A., Camara-Almirón, J., de Vicente, A. & Romero, D. Multifunctional amyloids in the biology of Gram-positive bacteria. Microorganisms 8, 2020 (2020).

    Article  Google Scholar 

  61. Schubeis, T. et al. Untangling a repetitive amyloid sequence: correlating biofilm-derived and segmentally labeled curli fimbriae by solid-state NMR spectroscopy. Angew. Chem. Int. Ed. 54, 14669–14672 (2015).

    Article  CAS  Google Scholar 

  62. Deshmukh, M., Evans, M. L. & Chapman, M. R. Amyloid by design: intrinsic regulation of microbial amyloid assembly. J. Mol. Biol. 12, 3631–3641 (2018).

    Article  Google Scholar 

  63. Cámara-Almirón, J., Caro-Astorga, J., de Vincente, A. & Romero, D. Beyond the expected: the structural and functional diversity of bacterial amyloids. Crit. Rev. Microbiol. 44, 653–666 (2018).

    Article  Google Scholar 

  64. Van Gerven, N., van der Verren, S. E., Reiter, D. M. & Remaut, H. The role of functional amyloids in bacterial virulence. J. Mol. Biol. 430, 3657–3684 (2018).

    Article  Google Scholar 

  65. Levkovich, S. A., Gazit, E. & Bar-Yosev, D. L. Two decades of studying functional amyloids in microorganisms. Trends Microbiol. 29, 251–265 (2020).

    Article  Google Scholar 

  66. Jain, N. & Chapman, M. R. Bacterial functional amyloids: order from disorder. BBA Prot. Proteom. 1867, 954–960 (2019).

    Article  CAS  Google Scholar 

  67. Otto, S. et al. Privatization of biofilm matrix in structurally heterogeneous biofilms. mSystems 5, e00425-20 (2020).

    Article  Google Scholar 

  68. Steinberg, N. et al. The extracellular matrix protein TasA is a developmental cue that maintains a motile subpopulation within Bacillus subtilis biofilms. Sci. Signal. 13, eaaw8905 (2020).

    Article  CAS  Google Scholar 

  69. Taglialegna, A. et al. Staphylococcal bap proteins build amyloid scaffold biofilm matrices in response to environmental signals. PLoS Pathog. 12, e1005711 (2016).

    Article  Google Scholar 

  70. Salinas, N., Povolotsky, T. L., Landau, M. & Kolodkin-Gal, H. Emerging roles of functional bacterial amyloids in gene regulation, toxicity, and immunomodulation. Microbiol. Mol. Biol. Rev. 85, e00062-20 (2021).

    Article  Google Scholar 

  71. Christensen, L. F. B. et al. The sheats of Methanospirillum are made of a new type of amyloid protein. Front. Microbiol. 9, 2729 (2018).

    Article  Google Scholar 

  72. Morris, R. J. et al. Natural variations in the biofilm-associated protein BslA from the genus. Bacillus. Sci. Rep. 7, 6730 (2017).

    Article  Google Scholar 

  73. Arnouteli, S., Bamford, N. C., Stanley-Wall, N. R. & Kovács, Á. T. Bacillus subtilis biofilm formation and social interactions. Nat. Rev. Microbiol. 19, 600–614 (2021).

    Article  Google Scholar 

  74. Kobayashi, K. & Iwano, M. BslA (YuaB) forms a hydrophobic layer on the surface of Bacillus subtilis biofilms. Mol. Microbiol. 85, 51–66 (2012).

    Article  CAS  Google Scholar 

  75. Hobley, L. A. et al. BslA is a self-assembling bacterial hydrophobin that coats the Bacillus subtilis biofilm. Proc. Natl Acad. Sci. USA 110, 33 (2013).

    Article  Google Scholar 

  76. Werb, M. et al. Surface topology affects wetting behavior of Bacillus subtilis biofilms. NPJ Biofilms Microbiomes 3, 11 (2017).

    Article  Google Scholar 

  77. Yu, H. Q. Molecular insights into extracellular polymeric substances in activated sludge. Environ. Sci. Microbiol. 54, 7742–7750 (2020).

    CAS  Google Scholar 

  78. Spaeth, R., Flemming, H.-C. & Wuertz, S. Sorption properties of biofilms. Water Sci. Technol. 37, 207–210 (1998).

    Article  Google Scholar 

  79. Schwartz, K., Ganesan, M., Payne, D. E., Solomon, M. J. & Boles, B. R. Extracellular DNA facilitates the formation of functional amyloids in Staphylococcus aureus biofilms. Mol. Microbiol. 99, 123–134 (2016).

    Article  CAS  Google Scholar 

  80. Okshevsky, M. & Meyer, R. L. The role of extracellular DNA in the establishment, maintenance and perpetuation of bacterial biofilms. Crit. Rev. Microbiol. 41, 341–352 (2015).

    Article  CAS  Google Scholar 

  81. Kästner, M. & Miltner, A. in The Future of Soil Carbon (eds Garcia, C., Nannipieri, P. & Hernandez, T.) 125–163 (Elsevier, 2018).

  82. Chrzanowski, L., Lawniczak, L. & Czaczyk, K. Why do microorganisms produce rhamnolipids? World J. Microbiol. Biotechnol. 28, 401–419 (2012).

    Article  CAS  Google Scholar 

  83. Aldeek, F. et al. Patterned hydrophobic domains in the exopolymer matrix of Shewanella oneidensis MR-1 biofilms. Appl. Environ. Microbiol. 79, 1400–1402 (2013).

    Article  CAS  Google Scholar 

  84. Campoccia, D., Montanaro, L. & Arciola, C. R. Extracellular DNA (eDNA). A major ubiquitous element of the bacterial biofilm architecture. Int. J. Mol. Sci. 22, 9100 (2021). This work presents a comprehensive overview of the fundamental structural role of eDNA and the contribution it offers to the complex architecture of the biofilm matrix by interaction with various other EPS components.

    Article  CAS  Google Scholar 

  85. de Aldecoa, A. L. I., Zafra, O. & González-Pastor, J. E. Mechanism and regulation of extracellular DNA release and its biological role in microbial communities. Front. Microbiol. 8, 1390 (2017).

    Article  Google Scholar 

  86. Panlilio, H. & Rice, C. V. The role of extracellular DNA in the formation, architecture, stability, and treatment of bacterial biofilms. Biotechnol. Bioeng. 118, 2129–2141 (2021). This work addresses, in particular, the role of eDNA for structure and stability of the EPS matrix and develops approaches to disrupt infectious biofilms.

    Article  CAS  Google Scholar 

  87. McDonough, E. K., Kamp, H. & Camili, A. Vibrio cholerae phosphatases required for the utilization of nucleotides and extracellular DNA as phosphate sources. Mol. Microbiol. 99, 453–469 (2016).

    Article  CAS  Google Scholar 

  88. Seviour, T. W. et al. The biofilm matrix scaffold of Pseudomonas aeruginosa contains G-quadruplex extracellular DNA structures. NPJ Biofilms Microbiomes 7, 27 (2021).

    Article  CAS  Google Scholar 

  89. Sørensen, S. J., Bailey, M., Hansen, L. H., Kroer, N. & Wuertz, S. Studying plasmid horizontal gene transfer in situ: a critical review. Nat. Rev. Microbiol. 3, 701–710 (2005).

    Article  Google Scholar 

  90. Devaraj, A. et al. The extracellular DNA lattice of bacterial biofilms is structurally related to Holliday junction recombination intermediates. Proc. Natl Acad. Sci. USA 116, 50 (2019). This work is the first paper to demonstrate that specific proteins normally associated with intracellular functions such as transcription and translation interact with eDNA outside the cell to provide biofilm structural stability.

    Article  Google Scholar 

  91. Buzzo, J. R. et al. Z-form extracellular DNA is a structural component of the bacterial biofilm matrix. Cell 184, 1–19 (2021).

    Article  Google Scholar 

  92. Randrianjatovo-Gbalou, I., Rouquette, P., Levebvre, D., Girbal-Neuhauser, E. & Marcato-Romain, C.-E. In situ analysis of Bacillus licheniformis biofilms: amyloid-like polymers and eDNA are involved in the adherence and aggregation of the extracellular matrix. Appl. Microbiol. 122, 1262–1274 (2017).

    Article  CAS  Google Scholar 

  93. Saxena, P., Joshi, Y., Rawat, K. & Bisht, R. Biofilms: architecture, resistance, quorum sensing and control mechanisms. Indian J. Microbiol. 59, 3–12 (2019).

    Article  Google Scholar 

  94. Dawson, L. F. et al. Extracellular DNA, cell surface protein and c-di-GMP promote biofilm formation in Clostidioides difficile. Sci. Rep. 11, 3244 (2021).

    Article  CAS  Google Scholar 

  95. Kavanaugh, J. S. et al. Identification of extracellular DNA-binding proteins in the biofilm matrix. mBio 10, e01137-19 (2019). This work discovers a series of lipoproteins with DNA-binding activity and develops an electrostatic net model in which membrane-attached lipoproteins function as anchor points between eDNA in the matrix and the bacterial cell surface.

    Article  Google Scholar 

  96. Jacubovics, N. S., Goodman, S. D., Mashburn-Warren, L., Stafford, G. P. & Cieplik, F. The dental plaque biofilm matrix. Periodontol 2000 86, 32–56 (2021).

    Article  Google Scholar 

  97. Chiba, A. et al. Staphylococcus aureus utilizes environmental RNA as a building material in specific polysaccharide-dependent biofilms. NPJ Biofilms Microbiomes 8, 17 (2022).

    Article  CAS  Google Scholar 

  98. Deatherage, B. L. & Cookson, B. T. Membrane vesicle release in bacteria, eukaryotes and archaea: a conserved yet underappreciated aspect of microbial life. Infect. Immun. 80, 1948–1957 (2012).

    Article  CAS  Google Scholar 

  99. Kikuchi, Y. et al. Diversity of physical properties of bacterial extracellular membrane vesicles revealed through atomic force microscopy imaging. Nanoscale 12, 7950–7959 (2020).

    Article  CAS  Google Scholar 

  100. Nagakubo, T., Nomura, N. & Toyofuku, M. Cracking open bacterial membrane vesicles. Front. Microbiol. 10, 3026 (2020).

    Article  Google Scholar 

  101. Schooling, S. R. & Beveridge, T. J. Membrane vesicles: an overlooked component in the matrices of biofilms. J. Bacteriol. 188, 5945–5957 (2006).

    Article  CAS  Google Scholar 

  102. Elhenawy, W., Debelyy, M. O. & Feldman, M. F. Preferential packing of acidic glycosidases and proteases into Bacteroides outer membrane vesicles. mBio 5, e00909-14 (2014).

    Article  Google Scholar 

  103. Brown, L., Wolf, J. M., Prados-Rosales, R. & Casadevall, A. Through the wall: extracellular vesicles in Gram-positive bacteria, mycobacteria and fungi. Nat. Rev. Microbiol. 13, 620–630 (2015).

    Article  CAS  Google Scholar 

  104. Liu, Y., Defourny, K. A. Y., Smid, E. J. & Abee, L. T. Gram-positive bacterial extracellular vesicles and their impact on health and disease. Front. Microbiol. 9, 1502 (2018).

    Article  CAS  Google Scholar 

  105. Toyofuku, M. Bacterial communication through membrane vesicles. Biosci. Biotechnol. Biochem. 83, 1599–1605 (2019).

    Article  CAS  Google Scholar 

  106. Morinaga, K., Yoshida, K., Takahashi, K., Nomura, N. & Toyofuku, M. Pecularities of biofilm formation by Paracoccus denitrificans. Appl. Microbiol. Biotechnol. 104, 2427–2433 (2020).

    Article  CAS  Google Scholar 

  107. Schwechheimer, C. & Kuehn, M. J. Outer-membrane vesicles from Gram-negative bacteria: biogenesis and functions. Nat. Rev. Microbiol. 13, 605–619 (2015).

    Article  CAS  Google Scholar 

  108. Baeza, N. & Mercade, E. Relationship between membrane vesicles, extracellular ATP and biofilm formation in Antarctic Gram-negative bacteria. Microb. Ecol. 81, 645–656 (2020).

    Article  Google Scholar 

  109. Park, M., Sun, Q., Liu, F., DeLisa, M. P. & Chen, W. Positional assembly of enzymes on bacterial outer membrane vesicles for cascade reactions. PLoS ONE 9, e97103 (2014).

    Article  Google Scholar 

  110. He, X. et al. Membrane vesicles are the dominant structural components of ceftazidime-induced biofilm formation in an oxacillin-sensitive MRSA. Front. Microbiol. 10, 571 (2019).

    Article  Google Scholar 

  111. Seike, S. et al. Outer membrane vesicles released from Aeromonas strains are involved in biofilm formation. Front. Microbiol. 11, 613650 (2021).

    Article  Google Scholar 

  112. Toyofuku, M. et al. Prophage-triggered membrane vesicle formation through peptoglycan damage in Bacillus subtilis. Nat. Comm. 8, 481 (2017).

    Article  Google Scholar 

  113. Piccolo, A. et al. in The Future of Soil Carbon (eds Garcia, C., Nainpieri, P. & Hernandez, T.) 87–124 (Elsevier, 2018).

  114. Jiao, N. et al. Microbial production of recalcitrant dissolved organic matter: long-term carbon storage in the global ocean. Nat. Rev. Microbiol. 8, 593–599 (2010).

    Article  CAS  Google Scholar 

  115. Zheng, T., Miltner, A., Liang, C., Nowak, K. M. & Kästner, M. Turnover of Gram-negative bacterial biomass-derived carbon through the microbial food web of an agricultural soil. Soil Biol. Biochem. 152, 108070 (2021).

    Article  CAS  Google Scholar 

  116. Liang, C., Amelung, W., Lehmann, J. & Kästner, M. Quantitative assessment of microbial necromass contribution to soil organic matter. Glob. Change Biol. 25, 3578–3590 (2019).

    Article  Google Scholar 

  117. Yang, J., Toyofuku, M., Sakai, R. & Nomura, N. Influence of the alginate production on cell-to-cell communication in Pseudomonas aeruginosa PAO1. Environ. Microbiol. Rep. 9, 239–249 (2017).

    Article  CAS  Google Scholar 

  118. Spitzer, J. From water and ions to crowded biomacromolecules: in vivo structuring of a prokaryotic cell. Microbiol. Mol. Biol. Rev. 75, 491–506 (2011). This work develops a theoretical model of molecular crowding in confined spaces as the base for enhanced multiple interactions among biomacromolecules, enabling multiple biochemical and physiological functions.

    Article  CAS  Google Scholar 

  119. Mittal, S., Chowhan, R. K. & Singh, L. R. Macromolecular crowding: friend or foe. Biochem. Biophys. Acta 1850, 1822–1831 (2015).

    Article  CAS  Google Scholar 

  120. Berk, V. et al. Molecular architecture and assembly principles of Vibrio cholerae biofilms. Science 337, 236–239 (2012).

    Article  CAS  Google Scholar 

  121. Fong, J. C. N. et al. Structural dynamics of RbmA governs plasticity of Vibrio cholerae biofilms. eLife 6, e26163 (2017).

    Article  Google Scholar 

  122. Peng, N. et al. The exopolysaccharide–eDNA interaction modulates 3D architecture of Bacillus subtilis biofilm. BMC Microbiol. 20, 111 (2020).

    Article  Google Scholar 

  123. Kanampalliwar, A. & Singh, D. V. Extracellular DNA builds and interacts with Vibrio polysaccharide in the biofilm matrix formed by Vibrio cholerae. Environ. Microbiol. Rep. 12, 594–606 (2020).

    Article  CAS  Google Scholar 

  124. Jennings, L. K. et al. Pel is a cationic exopolysaccharide that cross-links extracellular DNA in the Pseudomonas aeruginosa biofilm. Proc. Natl Acad. Sci. USA 112, 11353–11358 (2015).

    Article  CAS  Google Scholar 

  125. Irie, Y. et al. Self-produced exopolysaccharide is a signal that stimulates biofilm formation in Pseudomonas aeruginosa. Proc. Natl Acad. Sci. USA 109, 20632–20636 (2012).

    Article  CAS  Google Scholar 

  126. Yu, H. Y. et al. Elastase LasB of Pseudomonas aeruginosa promotes biofilm formation partly through rhamnolipid-mediated regulation. Can. J. Microbiol. 60, 227–235 (2014).

    Article  CAS  Google Scholar 

  127. Seviour, T. W. et al. Functional amyloids keep quorum sensing molecules in check. J. Biol. Chem. 290, 6457–6469 (2015). This paper shows how functional amyloids contain hydrophobic domains which bind signalling molecules with transient affinity, providing a pool of hydrophobic quorum sensing molecules.

    Article  CAS  Google Scholar 

  128. Grande, R. et al. Detection and physicochemical characterization of membrane vesicles (MVs) of Lactobacillus reuteri DSM 17938. Front. Microbiol. 8, 1040 (2017).

    Article  Google Scholar 

  129. Das, D. et al. Phenazine virulence factor binding to extracellular DNA is important for Pseudomonas aeruginosa biofilm formation. Sci. Rep. 5, 8398 (2015).

    Article  CAS  Google Scholar 

  130. Schiessl, K. et al. Phenazine production promotes antibiotic tolerance and metabolic heterogeneity in Pseudomonas aeruginosa biofilms. Nat. Commun. 10, 762 (2019).

    Article  CAS  Google Scholar 

  131. Saunders, S. H. et al. Extracellular DNA promotes efficient electron transfer by pyocyanin in Pseudomonas aeruginosa biofilms. Cell 182, 191–932 (2020).

    Article  Google Scholar 

  132. Zhang, Z. et al. Organic loading rate (OLR) regulation for enhancement of aerobic sludge granulation: role of key microorganism and their function. Sci. Tot. Environ. 653, 630–637 (2019).

    Article  CAS  Google Scholar 

  133. van Loosdrecht, M. C. M., Heijnen, J. J., Eberl, H., Kreft, J. & Picioreanu, C. Mathematical modelling of biofilm structures. Ant. Leeuwenhoek 81, 245–256 (2002).

    Article  Google Scholar 

  134. Fazli, M. et al. Regulation of biofilm formation in Pseudomonas and Burkholderia species. Env. Microbiol. 16, 1961–1981 (2014).

    Article  CAS  Google Scholar 

  135. Wolska, K., Grudniak, A. M., Rudnicka, Z. & Markowska, K. Genetic control of bacterial biofilms. J. Appl. Gen. 57, 225–238 (2016). This work gives a clear and comprehensive overview on quorum sensing molecules, c-di-GMP and small RNAs as regulators in the life cycle of some Gram-negative species.

    Article  CAS  Google Scholar 

  136. Poulin, M. B. & Kuperman, L. L. Regulation of biofilm exopolysaccharide production by cyclic di-guanosine monophosphate. Front. Microbiol. 12, 730980 (2021).

    Article  Google Scholar 

  137. Teschler, J. K. et al. Living in the matrix: assembly and control of Vibrio cholerae biofilms. Nat. Rev. Microbiol. 13, 255–268 (2015).

    Article  CAS  Google Scholar 

  138. Moormeier, D. E. & Bayles, K. W. Staphylococcus aureus biofilm: a complex developmental organism. Mol. Microbiol. 104, 365–367 (2017).

    Article  CAS  Google Scholar 

  139. Schilcher, K. & Horswill, A. R. Staphylococcal biofilm development: structure, regulation and treatment strategies. Microbiol. Mol. Biol. Rev. 84, e00026-19 (2020).

    Article  Google Scholar 

  140. Neu, T. R. & Lawrence, J. R. in Productive Biofilms (eds Muffler, K. & Ulber, R.) 1–51 (Springer International, 2014).

  141. Karampatzakis, A. et al. Measurement of oxygen concentrations in bacterial biofilms using transient state monitoring by single plane illumination microscopy. Biomed. Phys. Engin. Expr. 3, 035020 (2017).

    Article  Google Scholar 

  142. Wagner, M. & Horn, H. Optical coherence tomography in biofilm research: a comprehensive review. Biotechnol. Bioeng. 114, 1386–1402 (2017).

    Article  CAS  Google Scholar 

  143. Lawrence, J. R., Korber, D. R., Hoyle, B. D., Costerton, J. W. & Caldwell, D. E. Optical sectioning of microbial biofilms. J. Bacteriol. 173, 6558–6567 (1991).

    Article  CAS  Google Scholar 

  144. Lawrence, J. R., Swerhone, G. D. W., Kuhlicke, U. & Neu, T. R. In situ evidence for metabolic and chemical microdomains in the structured polymer matrix of bacterial microcolonies. FEMS Microbiol. Ecol. 92, fiw183 (2016).

    Article  Google Scholar 

  145. Lawrence, J. R., Swerhone, G. D. W. & Neu, T. R. Visualization of the sorption of nickel within exopolymer microdomains of bacterial microcolonies using confocal and scanning electron microscopy. Microbes Eniron 34, 76–82 (2019).

    Article  Google Scholar 

  146. Lawrence, J. R., Winkler, M. & Neu, T. R. Multi-parameter laser imaging reveals complex microscale biofilm matrix in a thick (4000 µm) aerobic methanol oxidizing community. Front. Microbiol. 9, 2186 (2018).

    Article  Google Scholar 

  147. Karwautz, C., Kus, G., Stöckl, M., Neu, T. R. & Lueders, T. Microbial megacities fueled by methane oxidation in a mineral spring cave. ISME J. 12, 87–100 (2018).

    Article  CAS  Google Scholar 

  148. Neu, T. R. & Kuhlicke, U. Fluorescence lectin bar-coding of glycoconjugates in the extracellular matrix of biofilm and bioaggregate forming microorganisms. Microorganisms 5, 5 (2017).

    Article  Google Scholar 

  149. Neu, T. R. & Lawrence, J. R. in Aquatic Biofilms: Ecology, Water Quality and Wastewater Treatment (eds Romani, A. M., Guasch, H. & Balaguer, M. D.) 29–45 (Caister Acad. Press, 2016).

  150. Raman, R., Raguraman, S., Venkataraman, G., Paulson, J. C. & Sasisekharan, R. Glycomics: an integrated system approach to structure–function relationships of glycans. Nat. Meth. 2, 817–824 (2005).

    Article  CAS  Google Scholar 

  151. Kellman, B. P. & Lewis, N. E. Big-data glycomics: tools to connect glycan biosynthesis to extracellular communication. Trends Biochem. Sci. 46, 284–300 (2021).

    Article  CAS  Google Scholar 

  152. Laughlin, S. T. & Bertozzi, C. R. Imaging the glycome. Proc. Natl Acad. Sci. USA 106, 12–17 (2009).

    Article  CAS  Google Scholar 

  153. Siegrist, M. S., Swarts, B. M., Fox, D. M., Lim, S. A. & Bertozzi, C. R. Illumination of growth, division and secretion by metabolic labeling of the bacterial cell surface. FEMS Microbiol. Rev. 39, 184–202 (2015).

    Article  Google Scholar 

  154. Geta-Zatorsky, N. et al. In vivo imaging and tracking of host–microbiota interactions via metabolic lableing of gut anaerobic bacteria. Nat. Med. 21, 1091–1100 (2015).

    Article  Google Scholar 

  155. Gregor, I. & Enderlein, J. Image scanning microscopy. Curr. Opin. Chem. l Biol. 51, 74–83 (2019).

    Article  CAS  Google Scholar 

  156. Kumar, A. et al. Dual-view plane illumination microscopy for rapid and spatially isotropic imaging. Nat. Protoc. 9, 2555–2573 (2014).

    Article  CAS  Google Scholar 

  157. Rooney, L. M., Amos, W. B., Hoskisson, P. A. & McConnell, G. Intra-colony channels in E. coli function as a nutrient uptake system. ISME J. 14, 2461–2473 (2020).

    Article  CAS  Google Scholar 

  158. Peterson, B. W. et al. Viscoelasticity of biofilms and their recalcitrance to mechanical and chemical challenges. FEMS Microbiol. Rev. 39, 234–245 (2015).

    Article  CAS  Google Scholar 

  159. Stewart, E., Ganesan, M., Younger, J. G. & Solomon, M. J. Artificial biofilms establish the role of matrix interactions in staphylococcal biofilm assembly and disassembly. Sci. Rep. 5, 13081 (2015).

    Article  CAS  Google Scholar 

  160. Gloag, E. S., Fabbri, S., Wozniak, D. J. & Stoodley, P. Biofilm mechanics: implications in infection and survival. Biofilm 2, 100017 (2020). This work describes very well how biofilm viscoelasticity due to the cohesion of multiple EPS components contributes to biofilm resilience to medical treatment and contributes to the virulence of chronic biofilm infections.

    Article  CAS  Google Scholar 

  161. Billings, N. et al. Material properties of biofilms — a review of methods for understanding permeability and mechanics. Rep. Progr. Phys. 78, 036601 (2015).

    Article  Google Scholar 

  162. Seviour, T. et al. The biofilm matrix scaffold of Pseudomonas aeruginosa contains G-quadruplex extracellular structures. NPJ Biofilms Microbiomes 7, 27 (2021).

    Article  CAS  Google Scholar 

  163. Díaz-Pascual, F. et al. Breakdown of Vibrio cholerae biofilm architecture induced by antibiotics disrupts community barrier function. Nat. Microbiol. 4, 2136–2145 (2019).

    Article  Google Scholar 

  164. Bergstrom, J. S. & Boyce, M. C. Mechanical behavior of particle filled elastomers. Rubber Chem. Technol. 72, 633–656 (1999).

    Article  CAS  Google Scholar 

  165. Qi, L. & Christopher, G. F. Rheological variability of Pseudomonas aeruginosa biofilms. Rheol. Acta 60, 219–230 (2021).

    Article  CAS  Google Scholar 

  166. Fabbri, S. et al. Fluid-driven interfacial instabilities and turbulence in bacterial biofilms. Environ. Microbiol. 19, 4417–4431 (2017).

    Article  CAS  Google Scholar 

  167. Risse-Buhl, U. et al. The role of hydrodynamics in shaping the composition and architecture of epilithic biofilms in fluvial ecosystems. Water Res. 127, 211–222 (2017).

    Article  CAS  Google Scholar 

  168. Picioreanu, C., Blauert, F., Horn, H. & Wagner, M. Determination of mechanical properties of biofilms by modelling the deformation measured using optical coherence tomography. Water Res. 145, 588–598 (2018).

    Article  CAS  Google Scholar 

  169. Prades, L. et al. Computational and experimental investigation of biofilm disruption dynamics induced by high-velocity gas jet impingement. mBio 11, 1 (2020).

    Article  Google Scholar 

  170. Fabbri, S. & Stoodley, P. in The Perfect Slime. Microbial Extracellular Polymeric Substances (EPS) (eds Flemming, H.-C., Neu, T. R. & Wingender, J.) 153–177 (IWA, 2017).

  171. Wilking, J. N., Angelini, T. E., Seminara, A., Brenner, M. P. & Weitz, D. A. Biofilms as complex fluids. MRS Bull. 36, 385–391 (2011).

    Article  CAS  Google Scholar 

  172. Cao, H., Habimana, O., Safari, A., Heffernan, R. & Casey, E. Revealing region-specific viscoelastic properties by means of a micro-rheological approach. NPJ Biofilms Microbiomes 2, 5 (2016).

    Article  Google Scholar 

  173. Saikat, J. et al. Nonlinear rheological characteristics of single species bacterial biofilms. NPJ Biofilms Microbiomes 6, 19 (2020).

    Article  Google Scholar 

  174. Charlton, S. G. V. et al. Regulating, measuring and modeling the viscoelasticity of bacterial biofilms. J. Bacteriol. 201, e00101-19 (2019).

    Article  CAS  Google Scholar 

  175. Pattem, J. et al. A multi-scale biophysical approach to develop structure–property relationships in oral biofilms. Sci. Rep. 8, 5691 (2018).

    Article  CAS  Google Scholar 

  176. Lieleg, O., Caldara, M., Baumgärtel, R. & Ribbeck, K. Mechanical robustness of Pseudomonas aeruginosa biofilms. Soft Matter 7, 3307–3314 (2011).

    Article  CAS  Google Scholar 

  177. Lei, W., Bruchmann, J., Rüping, J. L., Levkin, P. A. & Schwartz, T. Biofilm bridges forming structural networks on patterned lubricant-infused surfaces. Adv. Sci. 6, 1900519 (2019).

    Article  Google Scholar 

  178. Rozenbaum, R. T. et al. Role of viscoelasticity in bacterial killing by antimicrobials in differently grown Pseudomonas aeruginosa biofilms. Antimicrob. Agents Chemother. 63, e01972 (2019).

    Article  CAS  Google Scholar 

  179. Rahman, M. U. et al. Microrheology of Pseudomonas aeruginosa biofilms grown in wound beds. NPJ Biofilms Microbiomes 8, 49 (2022).

    Article  CAS  Google Scholar 

  180. Flemming, H.-C. et al. Who put the film in biofilm? The migration of a term from wastewater engineering to medicine and beyond. NPJ Biofilms Microbiomes 7, 10 (2021).

    Article  Google Scholar 

  181. Quan, K. et al. Water in bacterial biofilms: pores, channels, storage and transport functions. Crit. Rev. Microbiol. 48, 283–302 (2021).

    Article  Google Scholar 

  182. Katharios-Lanwermeyer, S. & O´Toole, G. A. Biofilm maintenance as an active process: evidence that biofilms work hard to stay put. J. Bacteriol. 204, e00587-21 (2022).

    Article  Google Scholar 

  183. Fritts, R. K., McCully, A. L. & McKinlay, J. B. Extracellular metabolism sets the table for microbial cross-feeding. Microbiol. Mol. Biol. Rev. 85, e00135-20 (2021).

    Article  Google Scholar 

  184. Arshad, Z. et al. Using stable isotope probing and fluorescence spectroscopy to examine the roles of substrate and soluble microbial products in extracellular polymeric substance formation in activated sludge processes. Sci. Tot. Environ. 178, 147875 (2021).

    Article  Google Scholar 

  185. Costa, O. Y. A., Pijl, A. & Kuramae, E. E. Dynamics of active potential bacterial and fungal interactions of acidobacterial EPS in soil. Soil Biol. Biochem. 148, 107916 (2020).

    Article  CAS  Google Scholar 

  186. Boutrif, M., Garel, M., Cortell, T. & Tamburini, C. Assimilation of marine extracellular polymeric substances by deep-sea prokaryotes in the NW Mediterranean Sea. Environ. Microbiol. Rep. 3, 705–709 (2011).

    Article  CAS  Google Scholar 

  187. Bharti, S. et al. Rv1717 is a cell wall-associated β-galactosidase of Mycobacterium tuberculosis that Is involved in biofilm dispersion. Front. Microbiol. 11, 611122 (2021).

    Article  Google Scholar 

  188. Wettstatt, S. Breaking free from home: biofilm dispersal by a glycosidase from Desulfovibrio vulgaris. Environ. Microbiol. 22, 557–558 (2020).

    Article  Google Scholar 

  189. Cherny, K. & Sauer, K. Untethering and degradation of the polysaccharide matrix are essential steps in the dispersion response of Pseudomonas aeruginosa biofilms. J. Bacteriol. 202, e00575-19 (2020).

    Article  Google Scholar 

  190. Pires, D. P., Oliveira, H., Melo, L. D. R., Sillancorva, S. & Azeredo, J. Bacteriophage-encoded depolymerases: their diversity and biotechnological applications. Appl. Microbiol. Biochechnol. 100, 2141–2151 (2016).

    CAS  Google Scholar 

  191. Lapébie, P., Lombard, V., Drula, E., Terrapon, N. & Henrissat, B. Bacteroidetes use thousands of enzyme combinations to break down glucans. Nat. Commun. 10, 2043 (2019).

    Article  Google Scholar 

  192. Glowacki, R. W. P. & Martens, E. C. If you eat it or secrete it, they will grow: the expanding list of nutrients utilized by the human gut bacteria. J. Bacteriol. 203, e00481-20 (2021).

    Article  Google Scholar 

  193. Larsbrink, J. & McKee, L. Bacteroidetes bacteria in the soil: glycan acquisition, enzyme secretion, and gliding motility. Adv. Appl. Microbiol. 110, 63–98 (2020).

    Article  CAS  Google Scholar 

  194. Brethauer, S., Shahab, R. & Studer, M. Impact of biofilms on the conversion of cellulose. Appl. Microbiol. Biotechnol. 104, 5201–5212 (2020).

    Article  CAS  Google Scholar 

  195. López-Mondéjar, R., Algora, C. & Baldrian, P. Lignocellulytic systems of soil bacteria: a vast and diverse toolbox for biotechnological conversion processes. Biotechnol. Adv. 37, 107374 (2019).

    Article  Google Scholar 

  196. Mitrofanova, O., Mardanova, A., Evtugyn, V., Bogomolnaya, L. & Sharipova, M. Effects of Bacillus serine proteases on the bacterial biofilms. Biomed. Res. Int. 2017, 8525912 (2017).

    Article  Google Scholar 

  197. Esoda, C. N. & Kuehn, M. J. Pseudomonas aeruginosa leucine aminopeptidase influences early biofilm composition and structure via vesicle-associated antibiofilm activity. mBio 10, e02548-19 (2019).

    Article  Google Scholar 

  198. Ningthoujam, S. et al. In vitro degradation of β-amyloid fibrils by microbial keratinase. Alzheimers Dement. (NY) 5, 154–163 (2019).

    Article  Google Scholar 

  199. Pressler, K. et al. Characterization of Vibrio cholerae’s extracellular nuclease Xds. Front. Microbiol. 10, 2057 (2019).

    Article  Google Scholar 

  200. Wasmund, K. et al. Genomic insights into diverse bacterial taxa that degrade extracellular DNA in marine sediments. Nat. Microbiol. 6, 885–898 (2021).

    Article  CAS  Google Scholar 

  201. Meylan, S., Andrews, I. W. & Collins, J. J. Targeting antibiotic tolerance, pathogen by pathogen. Cell 172, 1228–1238 (2018).

    Article  CAS  Google Scholar 

  202. Brauner, A., Fridman, O., Gefen, O. & Balaban, N. O. Distinguishing between resistance, tolerance and persistence to antibiotic treatment. Nat. Rev. Microbiol. 14, 320–330 (2016).

    Article  CAS  Google Scholar 

  203. Stewart, P. S. et al. Conceptual model of biofilm antibiotic tolerance that integrates phenomena of diffusion, metabolism, gene expression, and physiology. J. Bacteriol. 201, e00307-19 (2019).

    Article  CAS  Google Scholar 

  204. Doroshenko, N. et al. Extracellular DNA impedes the transport of vancomycin in Staphylococcus epidermidis biofilms preexposed to subinhibitory concentrations of vancomycin. Antimicrob. Agents Chemother. 58, 7273–7282 (2014).

    Article  Google Scholar 

  205. Tseng, B. S. et al. The extracellular matrix protects Pseudomonas aeruginosa biofilms by limiting the penetration of tobramycin. Environ. Microbiol. 15, 2865–2878 (2013).

    CAS  Google Scholar 

  206. Colvin, K. M. et al. The Pel polysaccharide can serve a structural and protective role in the biofilm matrix of Pseudomona aeruginosa. PLoS Pathog. 7, e1001264 (2011).

    Article  CAS  Google Scholar 

  207. Cerca, N., Jefferson, K. K., Oliveira, R., Pier, G. B. & Azeredo, J. Comparative antibody-mediated phagocytosis of Staphylococcus epidermidis cells grown in a biofilm or in the planktonic state. Infect. Immun. 74, 4849–4855 (2006).

    Article  CAS  Google Scholar 

  208. Jones, E. A., McGillivary, G. & Bakaletz, L. O. Extracellular DNA within a nontypeable Haemophilus influenzae-induced biofilm binds human β-defensin-3 and reduces its antimicrobial activity. J. Innate Immun. 5, 24–38 (2013).

    Article  CAS  Google Scholar 

  209. Wingender, J., Grobe, S., Fiedler, S. & Flemming, H.-C. in Biofilms in Aquatic Systems Vol. 242 (eds Keevil, C. W., Godfree, A. F., Holt, D. & Dow, C.) 93–100 (Royal Society of Chemistry, 1999).

  210. Hahn, M. M., González, J. F. & Gunn, J. S. Salmonella biofilms tolerate hydrogen peroxide by a combination of extracellular polymeric substance barrier function and catalase enzymes. Front. Cell. Infect. Microbiol. 11, 683081 (2021).

    Article  CAS  Google Scholar 

  211. Powell, L. et al. Quantifying the effects of antibiotic treatment on the extracellular polymer network of antimicrobial resistant and sensitive biofilms using multiple particle tracking. NPJ Biofilms Microbiomes 7, 1–11 (2021).

    Article  Google Scholar 

  212. Rosman, C. W., van der Mei, H. C. & Sjollema, J. Influence of sub-inhibitory concentrations of antimicrobials on micrococcal nuclease and biofilm formation in Staphylococcus aureus. Sci. Rep. 11, 13241 (2021).

    Article  CAS  Google Scholar 

  213. Ranieri, M. R. M., Whitchurch, C. M. & Burrows, L. L. Mechanisms of biofilm stimulation by subinhibitory concentrations of antimicrobials. Curr. Opin. Microbiol. 45, 164–169 (2018).

    Article  CAS  Google Scholar 

  214. Lin, J., Wang, Z., Zang, Y., Zhang, D. & Xin, Q. Detection of respiration changes inside biofilms with microelectrodes during exposure to antibiotics. J. Environ. Sci. Health A Tox. Hazard. Subst. Environ. Eng. 54, 202–207 (2019).

    Article  CAS  Google Scholar 

  215. Narten, M., Rosin, N., Schobert, M. & Tielen, P. Susceptibility of Pseudomonas aeruginosa urinary tract isolates and influence of urinary tract conditions on antibiotic tolerance. Curr. Microbiol. 64, 7–16 (2012).

    Article  CAS  Google Scholar 

  216. von Ohle, C. et al. Real-time microsensor measurement of local metabolic activities in ex vivo dental biofilms exposed to sucrose and treated with chlorhexidine. Appl. Environ. Microbiol. 76, 2326–2334 (2010).

    Article  Google Scholar 

  217. Bui, L. M. G., Conlon, B. P. & Kidd, S. P. Antibiotic tolerance and the alternative lifestyles of Staphylococcus aureus. Essays Biochem. 61, 71–79 (2017).

    Article  Google Scholar 

  218. Ciofu, O., Moser, C., Jensen, P. Ø. & Høiby, N. Tolerance and resistance of microbial biofilms. Nat. Rev. Microbiol. https://doi.org/10.1038/s41579-022-00682-4 (2022).

    Article  Google Scholar 

  219. Dworkin, J. & Shaw, I. M. Exit from dormancy in microbial organisms. Nat. Rev. Microbiol. 8, 890–896 (2010).

    Article  CAS  Google Scholar 

  220. Yan, J. & Bassler, B. L. Surviving as a community: antibiotic tolerance and persistence in bacterial biofilms. Cell Host Microbe 26, 15–21 (2019).

    Article  CAS  Google Scholar 

  221. Sindeldecker, D. et al. Novel aminoglycoside-tolerant phoenix colony variants of Pseudomonas aeruginosa. Antimicrob. Agents Chemother. 64, e00623-20 (2020).

    Article  Google Scholar 

  222. Abe, K., Nomura, N. & Suzuki, S. Biofilms: hot spots of horizontal gene transfer (HGT) in aquatic environments, with a focus on a new HGT mechanism. FEMS Microbiol. Ecol. 96, fiaa031 (2020).

    Article  CAS  Google Scholar 

Download references

Acknowledgements

The authors are grateful to I. C. H. Tan and C. Mayer for help with designing the drafts for Figs. 2 and 5a, respectively.

Author information

Authors and Affiliations

Authors

Contributions

H.-C.F., T.R.N., P.H.N., T.S. and P.S. researched data for the article. H.-C.F., E.D.v.H., T.R.N., P.H.N., T.S., P.S., J.W. and S.W. contributed substantially to discussion of the content. H.-C.F., T.R.N., P.H.N. and P.S. wrote the article. H.-C.F., T.R.N., J.W. and P.S. reviewed and/or edited the manuscript before submission. H.-C.F. assembled the team and coordinated the writing process.

Corresponding author

Correspondence to Hans-Curt Flemming.

Ethics declarations

Competing interests

The authors declare no competing interests.

Peer review

Peer review information

Nature Reviews Microbiology thanks Kendra Rumbaugh and the other, anonymous, reviewer(s) for their contribution to the peer review of this work.

Additional information

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Supplementary information

Glossary

Extracellular polymeric substances

(EPS). Microbial biopolymers such as polysaccharides, proteins, extracellular DNA (eDNA) and others, forming the biofilm matrix.

Extracellular matrix

The non-cellular component present within all tissues and organs, sometimes also used as an alternative to the term extracellular polymeric substances (EPS); frequently used in the medical context.

Amyloids

Aggregates of proteins in fibrillar morphology. Pathogenic amyloids form by misfolding of previously normal structures. In biofilms, amyloids fulfil many functions, including, for example, matrix stabilization, nutrient storage, desiccation resistance and others.

Humic-like microbially derived refractory substances

Remains of bacterial cells that are not readily degraded after cell death. As high molecular weight compounds they remain present within microbial communities, contributing to the polymeric matrix.

Humic substances

The organic components of humus. Humic substances are hetero-polycondensates based on a motif of aromatic nuclei with phenolic and carboxylic substituents linked together. They can form aggregates, provide cation complexation sites and regulate the bioavailability of metal ions.

Transparent exopolymer particles

Extracellular acidic polysaccharides produced by phytoplankton and bacteria in saltwater, freshwater and wastewater; they are extremely abundant and play a significant role in biogeochemical cycling of carbon and other elements in water.

Collective biological systems

Systems such as forests, beehives, coral reefs or kelp fields that show emerging properties which exceed those of the sum of the single individuals. Also known as ‘extended organisms’.

Psl

An extracellular polysaccharide of Pseudomonas aeruginosa and an important structural and functional feature of P. aeruginosa biofilms. Psl is rich in galactose and mannose.

Pel

An extracellular polysaccharide of Pseudomonas aeruginosa and an important structural and functional feature of P. aeruginosa biofilms. The structure of Pel is not fully characterized but it is a cationic polysaccharide, differing from Psl and alginate.

Ecotin

A protease inhibitor, formed in the periplasmic space of Gram-negative bacteria, inhibiting neutrophil elastase.

Anammox

(Ammonium oxidation). The reaction of nitrite and ammonium ions leading directly to dinitrogen and water.

Rhamnolipids

Amphiphilic glycolipids consisting of a monosaccharide or disaccharide connected by a glycosidic bond to a fatty acid; they act in various roles in the EPS matrix.

Necromass

The mass of dead biological material, including microorganisms.

Phenazine

The chemical description of the class of dibenzo annulated pyrazine; it embraces pyocyanine as a subclass in which one of the nitrogen atoms is substituted with a methyl group.

Glycomics

The study of all glycan structures in biology and a subset of glycobiology. Glycomics focuses on the identification of structure and function of the total collection of glycans (the glycome) produced by biological systems under specified conditions of time, space and environment.

Mesolens

A novel microscope objective lens that combines a high numerical aperture with a large field of view of up to 6 mm combined with high spatial resolution.

Nanoscopy

A term describing light microscopy techniques at a resolution across the diffraction limit of light. The techniques include localization or blink microscopy, stimulated emission depletion microscopy and, more recently, MinFlux. By exploiting switchable fluorochromes, achieving a resolution of 20–10 nm down to a few nanometres becomes possible.

Persisters

A subpopulation of transiently antibiotic-tolerant bacterial cells that are often slow growing or growth arrested, and are able to resume growth after a lethal stress.

Phoenix phenotypes

Phoenix colonies that grow out of the zone of clearance of antibiotic-loaded beads from lawn biofilms while there are still very high concentrations of antibiotic present, suggesting an antibiotic-resistant phenotype.

Rights and permissions

Springer Nature or its licensor holds exclusive rights to this article under a publishing agreement with the author(s) or other rightsholder(s); author self-archiving of the accepted manuscript version of this article is solely governed by the terms of such publishing agreement and applicable law.

Reprints and permissions

About this article

Check for updates. Verify currency and authenticity via CrossMark

Cite this article

Flemming, HC., van Hullebusch, E.D., Neu, T.R. et al. The biofilm matrix: multitasking in a shared space. Nat Rev Microbiol 21, 70–86 (2023). https://doi.org/10.1038/s41579-022-00791-0

Download citation

  • Accepted:

  • Published:

  • Issue Date:

  • DOI: https://doi.org/10.1038/s41579-022-00791-0

This article is cited by

Search

Quick links

Nature Briefing Microbiology

Sign up for the Nature Briefing: Microbiology newsletter — what matters in microbiology research, free to your inbox weekly.

Get the most important science stories of the day, free in your inbox. Sign up for Nature Briefing: Microbiology